Figures
Abstract
Insects use their sensitive and selective olfactory system to detect outside chemical odorants, such as female sex pheromones and host plant volatiles. Several groups of olfactory proteins participate in the odorant detection process, including odorant binding proteins (OBPs), chemosensory proteins (CSPs), odorant receptors (ORs), ionotropic receptors (IRs) and sensory neuron membrane proteins (SNMPs). The identification and functional characterization of these olfactory proteins will enhance our knowledge of the molecular basis of insect chemoreception. In this study, we report the identification and differential expression profiles of these olfactory genes in the black cutworm moth Agrotis ipsilon. In total, 33 OBPs, 12 CSPs, 42 ORs, 24 IRs, 2 SNMPs and 1 gustatory receptor (GR) were annotated from the A. ipsilon antennal transcriptomes, and further RT-PCR and RT-qPCR revealed that 22 OBPs, 3 CSPs, 35 ORs, 14 IRs and the 2 SNMPs are uniquely or primarily expressed in the male and female antennae. Furthermore, one OBP (AipsOBP6) and one CSP (AipsCSP2) were exclusively expressed in the female sex pheromone gland. These antennae-enriched OBPs, CSPs, ORs, IRs and SNMPs were suggested to be responsible for pheromone and general odorant detection and thus could be meaningful target genes for us to study their biological functions in vivo and in vitro.
Citation: Gu S-H, Sun L, Yang R-N, Wu K-M, Guo Y-Y, Li X-C, et al. (2014) Molecular Characterization and Differential Expression of Olfactory Genes in the Antennae of the Black Cutworm Moth Agrotis ipsilon. PLoS ONE 9(8): e103420. https://doi.org/10.1371/journal.pone.0103420
Editor: Richard David Newcomb, Plant and Food Research, New Zealand
Received: March 20, 2014; Accepted: June 28, 2014; Published: August 1, 2014
Copyright: © 2014 Gu et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: The authors confirm that all data underlying the findings are fully available without restriction. All relevant data are within the paper and its Supporting Information files.
Funding: This work was supported by the China National 973? Basic Research Program (Grant No. 2012CB114104) the National Natural Science Foundation of China (Grant No. 31071694, 31171858 and 31272048) and the Foundation for Innovative Research Groups of the National Natural Science Foundation of China (Grant No. 31321004). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors declare that they have no competing interests.
Introduction
Insects use their sensitive and selective antennae, which express various olfactory proteins, to detect air borne odorant molecules, such as sex pheromones and plant volatiles. Species-specific pheromone molecules and general plant volatiles enter the sensillum lymph of the different types of antennae sensilla via the multipores of the insect cuticle [1], [2]. During the last 30 years, our knowledge of the molecular and cellular basis of insect chemoreception has greatly expanded. It is commonly accepted that several different groups of antennae-enriched olfactory proteins participate in the first stage of the detection of olfactory signals, including odorant binding proteins (OBPs), chemosensory proteins (CSPs), odorant receptors (ORs), ionotropic receptors (IRs) and sensory neuron membrane proteins (SNMPs) [3].
Insect OBPs are small water-soluble olfactory proteins that are presumed to be synthesized by non-neuronal auxiliary cells (trichogen and tormogen cells) of the sensory neurons and secreted into the sensillum lymph in high concentrations (up to 10 mM) [4]–[7]. The insect OBPs are commonly believed act as carrier proteins to transport odorants to the olfactory receptors. Functional studies of insect OBPs at both molecular and behavior levels have proven that insect OBPs are indispensable in insect chemoreception. For example, Drosophila OBP LUSH is required for the activation of pheromone-sensitive chemosensory neurons by the pheromone 11-cis vaccenyl acetate (cVA) [8], [9]. Additionally, in the fire ant Solenopsis invicta, the pheromone binding protein gene Gp-9 regulates the colony social organization between the monogyne social form (with a single queen) and the polygyne form (with multiple queens) [10].
Insect CSPs, which were also called OS-D like proteins [11] or sensory appendage proteins (SAPs) [12], represent one novel group of olfactory proteins that are involved in insect olfaction. These proteins have shown broad expression profiles in chemosensory tissues, including antennae [13]–[17], maxillary palps [18], labial palps [18], [19] and proboscis [20]. However, these proteins are also found in non-chemosensory organs, such as legs [21], [22], wings [23], [24] and pheromone glands [15]. Functional studies of insect CSPs revealed that these proteins have multiple-functions in insect chemoreception, growth and development. For example, in the tsetse fly Glossina morsitans morsitans, the female antennae-enriched CSP transcripts were showed remarkable expression levels after a blood meal, which suggested that these proteins participate in the female host-seeking behavior [14]. In the American cockroach Periplaneta americana, one CSP homologous gene named P10 was expressed 30 times higher in regenerating legs than in normal legs, which indicated that the P10 gene had a putative function in the regeneration of insect legs [21], [22]. In the migratory locust Locusta migratoria, the antennae-expressed CSP gene has been proposed to regulate the rapid switch between attraction and repulsion behaviors [25].
The insect odorant receptors (ORs) are odorant-gated ion channels which composed of one odorant-binding subunit and the olfactory coreceptor Orco [26], [27]. The functional study of insect ORs, particularly the pheromone receptors (PRs), revealed their essential role in insect olfaction [28], [29]. The classical method to identify and annotate insect OR genes is through bioinformatic screenings of genomic sequences. At present, using this method, insect OR genes have been identified and annotated from various insect species, including Drosophila melanogaster [30]–[32], Anopheles gambiae [33], Aedes aegypti [34], Apis mellifera [35], Nasonia vitripennis [36], Bombyx mori [37], Tribolium castaneum [38], and Acyrthosiphon pisum [39].
Recently, a novel chemosensory receptor family called ionotropic receptors (IRs) was discovered in D. melanogaster [40]. In total, 66 IRs, which included two putative conserved coreceptors, IR25a and IR8a, were identified by screening D. melanogaster genomic data [41]. The expression analysis revealed that 15 DmelIR genes were specially expressed in the antennae [40]. The misexpression of DmelIR84a and DmelIR92a conferred ectopic olfactory responses to the electrophysiology-activated compounds phenylacetaldehyde and ammonia, respectively [40]. Thus far, different IR genes have been identified and annotated in various insect species, including D. melanogaster [40], B. mori [41], Spodoptera littoralis [42], A. gambiae [43], Manduca sexta [44], Cydia pomonella [45], and Helicoverpa armigera [46].
Previously, functional studies of insect olfactory genes primarily focused on model species, such as D. melanogaster and B. mori, whose genomic data are available. However, the functional studies of olfactory genes of other insect species have been restricted due the deficiency of the genomic data for these species. Recently, the high-throughput sequencing of antennae and other tissues have proved to be an efficient strategy for identifying and annotating different types of olfactory genes in various insect species, including A. gambiae [43], M. sexta [44], C. pomonella [45], H. armigera [46], Cotesia vestalis [47], Agrilus planipennis [48], Aphis gossypii [49], S. littoralis [50], Ips typographus and Dendroctonus ponderosae [51].
In the present study, using a next-generation sequencing (NGS) 454 GS FLX platform, we have identified and annotate several families of chemosensory genes (including OBPs, CSPs, ORs, IRs and SNMPs) from the antennae of the black cutworm moth Agrotis ipsilon (Hufnagel) (Lepidoptera: Noctuidae), which is known as a destructive pest of many crops [52]–[53]. Using semi-quantitative RT-PCR and real-time quantitative-PCR (RT-qPCR), we have screened a number of antennae-specific or enriched olfactory genes from the A. ipsilon antennal transcriptomes, which may play important functions in the chemoreception of A. ipsilon.
Results and Discussion
454 sequencing and de novo assembly
Two non-normalized cDNA libraries of the male and female A. ipsilon antennae were constructed. After a single sequencing run using the 454 GS FLX platform, a total of 551388 (mean length 539 bp) and 537572 raw reads (mean length 548 bp) were produced from the male and female antennae samples, respectively. After trimming adaptor sequences, contaminating sequences and low quality sequences, 550456 (mean length 531 bp) and 536474 clean reads (mean length 540 bp) from male and female antennae, respectively, remained for the following assembly.
All clean reads from male and female antennae were assembled and produced 40126 (mean length 1072 bp) and 41358 (mean length 1054 bp) unigenes, respectively. Furthermore, we assembled all clean reads from male and female antennae together and finally generated 48795 unigenes. Among these unigenes, 41173 are contigs (84.4%) and 7622 are singletons (15.6%). The assembled unigene lengths ranged from 100 bp to 15432 bp, with an average length of 967 bp. The size distribution of the assembled unigenes is shown in Figure 1. An overview of the sequencing and assembly process is presented in Table 1.
Homology searching of A. ipsilon antennal unigenes with other insect species
We search for homologs in other insect species using the BLASTx and BLASTn programs with the e-value cut-off of 10e-5 [54]. The results indicated that 25180 of the 48795 unigenes (51.6%) had BLASTx hits in the non-redundant protein (nr) databases and that 17947 unigenes (36.8%) had BLASTn hits in the non-redundant nucleotide sequence (nt) databases. Some unigenes are homologous to more than one species. Most annotated A. ipsilon antennal unigenes have the best hits with Lepidoptera insect genes (8542 of the 17947 nt-hit unigenes); the highest hits included 2818 unigenes that were homologous to B. mori genes, 1820 unigenes that were homologous to H. armigera genes. The second highest hits are with Dipteran species genes, with 276 hits of D. melanogaster genes, and 392 and 383 hits that were homologous to genes of the mosquitoes A. gambiae and A. aegypti, respectively. The other unigenes were found to be homologous to genes from the wasp N. vitripennis (348 hits), the beetle T. castaneum (244 hits) and from the western honey bee A. mellifera (261 hits) (Figure 2).
All A. ipsilon antennal unigenes were used in BLASTn to search the GenBank entries. The best hits with an E-value < = 1.0E-5 for each query were grouped according to species.
Functional annotation of the A. ipsilon antennal unigenes
Similar to those genes that were found in the antennal transcriptomes of M. sexta [44], S. littoralis [55] and H. armigera [46], most A. ipsilon antennal unigenes (approximately 72%) could not be assigned to a Gene Ontology (GO) category. In total, 11987 male antennal unigenes and 12240 female antennal unigenes were annotated into different functional groups (biological process, cellular components and molecular functions) according to GO analysis [56] (Figure 3). Some transcripts were annotated into more than one GO category. The numbers of each GO category were similar between the male and female antennal transcriptomes (Figure 3). The cellular process (6301 male antennal unigenes and 6425 female antennal unigenes) and metabolic process (5243 male antennal unigenes and 5349 female antennal unigenes) GO categories were most abundantly represented within the biological process GO ontology. In the cellular components GO ontology, the transcripts were primarily distributed in the cell (7148 male antennal unigenes and 7308 female antennal unigenes) and in cell part (6619 male antennal unigenes and 6752 female antennal unigenes). The GO analysis also showed that the binding (4705 male antennal unigenes and 4787 female antennal unigenes) and catalytic activity (5133 male antennal unigenes and 5210 female antennal unigenes) were most abundant in the molecular function ontology (Figure 3).
Chemosensory genes are highly abundant in the A. ipsilon antennae
Because a non-normalized cDNA library was used for 454 sequencing in this study, the number of reads of per unigene can represent the relative mRNA abundance in the A. ipsilon antennal transcriptomes. Among the top 500 most highly abundant transcripts, 89 transcripts are annotated as olfactory genes, which suggests their involvement in insect chemosensory reception, including olfactory receptors, odorant-binding proteins, chemosensory proteins, antennal cytochrome P450s, antennal-enriched UDP-glycosyltransferases, antennal oxidoreductases, antennal aldehyde oxidases, sensory neuron membrane proteins and takeout-like proteins (Table S1).
Candidate odorant binding proteins in the A. ipsilon antennae
OBPs are believed to be involved in the initial biochemical recognition steps in insect odorant perception by capturing and transporting odorant molecules to the olfactory receptors (ORs) [57]–[59]. In the A. ipsilon antennal transcriptomes, a total of 33 OBP genes were annotated (Table 2) based on the tBLASTn results. The number of A. ipsilon OBP identified in present study is a little fewer than the number identified from the genome of B. mori (44) [60], A. gambiae (57) [61] and D. melanogaster (51) [62], so there may still some OBP genes are not identified from the A. ipsilon antennae due to their low expression level. Among the identified 33 OBP genes, 28 have intact ORFs with lengths ranging from 402 bp to 759 bp. The RPKM value analysis revealed that 9 OBP genes (PBP1, PBP2, PBP3, GOBP1, GOBP2, OBP4, OBP11, OBP18 and OBP24) are highly abundant in the male and female antennal transcriptomes (RPKM>1000) (Table 2). The RT-PCR results indicated that 22 OBP genes (PBP1, PBP2, PBP3, GOBP1, GOBP2, OBP1, OBP2, OBP4, OBP5, OBP9, OBP11, OBP12, OBP13, OBP15, OBP16, OBP17, OBP19, OBP20, OBP21, OBP22, OBP24 and OBP26) are uniquely or primarily expressed in the male and female antennae (Figure 4). Based on the different expression profiles of these OBPs in male and female antennae, we suggest these male antennae-enriched expressed OBPs are involved in sex pheromone detection, whereas female antennae-enriched expressed OBPs play important roles in locating suitable host plants and oviposition sites.
MA: male antennae; FA: female antennae; Bo: body. Pheromone gland rather than body was used in the analysis of AipsOBP6 and AipsCSP2. Antennae specific or enriched genes are labeled with a red pentagram. β-actin was used as an internal reference gene to test the integrity of each cDNA template; the similar intensity of β-actin bands among different tissues indicates the use of equal template concentrations.
Furthermore, real-time quantitative PCR (RT-qPCR) analysis was performed to compare the accurate quantitative expression levels of these OBP genes among different tissues between sexes, and the results suggested that the three PBP genes (PBP1, PBP2 and PBP3) are expressed higher in the male antennae than in the female antennae (p<0.01) (Figure 5). However, the RT-qPCR results lack concordance with the RPKM values, this reason may be the sequencing depth of 454 is not good enough. PBP1 and PBP2 showed high binding affinities with the two main sex pheromones of A. ipsilon, whereas PBP3 specifically binds to the minor amount sex pheromone Z11-16: Ac with a high binding ability [63]. In contrast, the expression levels of GOBP1, GOBP2 and OBP17 were much higher in the female antennae than in the male antennae (Figure 5). Interestingly, one OBP (OBP6) was primarily expressed in the pheromone gland (PG) (Figure 4 and Figure 5); this result was also reported in another study [64]. Unlike the common antennae-enriched OBPs, this PG-expressed OBP may play a different role in odorant and pheromone detection and transportation.
MA: male antennae; FA: female antennae; Bo: body. Pheromone gland rather than body was used in the analysis of AipsOBP6. The internal controls β-actin and ribosomal protein S3 were used to normalize transcript levels in each sample. This figure was presented using β-actin as the reference gene to normalize the target gene expression and to correct sample-to-sample variation; similar results were obtained with ribosomal protein S3 as the reference gene. The standard error is represented by the error bar, and the different letters (a, b, c) above each bar denote significant differences (p<0.05).
Candidate chemosensory proteins in the A. ipsilon antennae
Chemosensory proteins (CSPs) represent a new class of soluble carrier proteins in the lymph of insect antennal chemosensilla and they are proposed to play similar functions as OBPs in insect chemoreception [65]. In this study, we have identified 12 novel CSP genes in the A. ipsilon antennae (Table 3). Based on the extensive expression profiles of CSPs, the remaining CSPs which expressed in other tissues such as legs and wings may not be identified in present study. In total, 11 of the novel genes had intact ORFs, and the protein sequences had the typical four conserved cysteines, which are recognized as the signature feature of insect CSPs [65]. The RPKM value analysis revealed that 4 CSP genes (CSP4, CSP7, CSP9 and CSP10) are highly abundant in the male and female antennal transcriptomes (RPKM>1000) (Table 3). The RT-PCR and RT-qPCR results indicated that 3 CSP genes (CSP8, CSP9 and CSP10) are highly expressed in the male and female antennae (Figure 4 and Figure 6). This result suggested that these three antennae-enriched CSPs might play essential roles in the chemical communication process in insects. Interestingly, one CSP gene (CSP2) was not expressed in the antennae but was specifically expressed in the female pheromone gland (PG) (Figure 4 and Figure 6). CSPs that are expressed in the pheromone gland of the cabbage armyworm M. brassicae can bind sex pheromone analogs, which suggests that these CSPs may play a role in pheromone capture [15]. In Heliothis virescens and B. mori, CSPs are all detected in the pheromone gland [66]–[68]. This observation suggests the possible involvement of these proteins in carrying and releasing sex pheromones, as demonstrated for the antennal OBPs and CSPs. The insect may use these female PG-enriched OBPs and CSPs to auto-detect and monitor the sex pheromones released by themselves [69]–[70].
MA: male antennae; FA: female antennae; Bo: body. Pheromone gland rather than body was used in the analysis of AipsCSP2. The internal controls β-actin and ribosomal protein S3 were used to normalize transcript levels in each sample. This figure was presented using β-actin as reference gene to normalize the target gene expression and correct sample-to-sample variation; similar results were obtained with ribosomal protein S3 as the reference gene. The standard error is represented by the error bar, and the different letters (a, b) above each bar denote significant differences (p<0.05).
Candidate olfactory receptors in the A. ipsilon antennae
Insect olfactory receptors (ORs) are the most important players in sex pheromone and general odorant detection. In the present study, we have identified 42 OR genes (41 typical ORs and one atypical coreceptor) from the A. ipsilon antennal transcriptomes (Table 4). In insect, the axons from the sensory neurons converge into glomeruli in the antennal lobe. There are 66 glomeruli in the antennae lobe of the male A. ipsilon moth [71], based on the hypothesis that the number of the glomeruli equals the number of olfactory receptors [72], [73], we predict there are about 24 OR genes still need to be identified. In total, 12 of the 42 ORs have intact ORFs. The RPKM value analysis revealed that the ORco had the highest expression level among the 42 ORs, with RPKM value of 741 and 997 in the male and female antennae, respectively. The other 41 typical ORs, however, showed a relative low expression level (RPKM ranged from 0 to 567) compared with the ORco, OBP and CSP genes. Three ORs (OR1, OR3 and OR4) showed a higher RPKM in the male antennae than in the female antennae (more than 20 times) (Table 4). The RT-PCR and RT-qPCR results indicated that 35 ORs were exclusively or primarily expressed in the antennae. Among these ORs, 4 ORs (OR1, OR3, OR4 and OR14) have male antennae-specific expression (Figure 7 and Figure 8), which suggests that these ORs may play essential roles in the detection of sex pheromones. In total, 4 ORs (OR6, OR7, OR8 and OR23) have female antennae-enriched expression (Figure 7 and Figure 8), which suggests that these ORs may play important roles in the detection of general odorants, such as host plant volatiles. The OR tree from three Lepidoptera insects are extremely divergent; however, the olfactory coreceptor family and the pheromone receptor family are highly conserved (Figure 9).
MA: male antennae; FA: female antennae; Bo: body. Genes that are equally expressed in the male and female antennae are labeled with a red pentagram. Genes that are specifically or primarily expressed in the male antennae are labeled with a red circle. Genes that are specifically or primarily expressed in the female antennae are labeled with a red triangle. β-actin was used as the internal reference gene to test the integrity of each cDNA templates; the similar intensity of β-actin bands among different tissues indicates the use of equal template concentrations.
MA: male antennae; FA: female antennae; Bo: body. The internal controls β-actin and ribosomal protein S3 were used to normalize transcript levels in each sample. This figure was presented using β-actin as the reference gene to normalize the target gene expression and to correct sample-to-sample variation; similar results were obtained with ribosomal protein S3 as the reference gene. The standard error is represented by the error bar, and the different letters (a, b, c) above each bar denote significant differences (p<0.05).
The protein names and sequences of ORs that were used in this analysis are listed in Table S5.
Candidate ionotropic receptors in the A. ipsilon antennae
Insect chemosensory ionotropic receptors (IRs) belong to an ancient chemosensory receptor family, that was first discovered in D. melanogaster and are expressed in sensory neurons that respond to different odorants but that do not express either ORs or gustatory receptors (GRs) [40]. The misexpression of D. melanogaster IRs conferred ectopic odorant responsiveness [40]. At present, 66 IRs in D. melanogaster [41], 12 IRs in the noctuid S. littoralis [42], 15 IRs in C. pomonella [45] and 12 IRs in H. armigera [46] have been identified. In the present study, we have identified 24 IRs, including two highly conserved coreceptors, IR8a and IR25a, from the A. ipsilon antennal transcriptomes (Table 5). Five of the IR genes, including coreceptors IR8a and IR25a, had intact ORFs. Eighteen of these 24 IRs showed high amino acid identity (52%–90%) with three Lepidoptera insects, C. pomonella, S. littoralis and B. mori. Similar to the ORs, the RPKM value analysis revealed that all the 24 IRs showed a relative low expression level (RPKM value ranged from 0 to 69) compared with the OBPs and CSPs. The antennae-enriched IRs may play important roles in odorant detection; 15 D. melanogaster IRs [40], 10 H. armigera IRs [46] and 7 S. littoralis IRs [42] were expressed exclusively in the antennae. Our RT-PCR and RT-qPCR results indicated that 14 A. ipsilon IRs (IR8a, IR25a, IR21a, IR41a, IR75q.1, IR75q.2, IR76b, IR87a, IR1, IR3, IR4, IR8, IR12 and IR13) are highly expressed in the antennae; in particular, one IR IR12 was specifically expressed in the male antennae (Figure 7 and Figure 10), which suggested that this IR may be devoted to the response to the female sex pheromones. IRs from different insect species are extremely divergent; however, the two coreceptors IR8a and IR25a are highly conserved among different insect species (Figure 11).
MA: male antennae; FA: female antennae; Bo: body. The internal controls β-actin and ribosomal protein S3 were used to normalize transcript levels in each sample. This figure was presented using β-actin as the reference gene to normalize the target gene expression and to correct sample-to-sample variation; similar results were obtained with ribosomal protein S3 as the reference gene. The standard error is represented by the error bar, and the different letters (a, b, c) above each bar denote significant differences (p<0.05).
The protein names and sequences of IRs that were used in this analysis are listed in Table S6.
Candidate sensory neuron membrane proteins and gustatory receptors in the A. ipsilon antennae
Insect SNMPs are two trans-membrane domain-containing proteins that are suggested to play significant roles in insect chemoreception [74]–[76]. Two SNMP subfamilies, SNMP1 and SNMP2, were identified in insects; however, these subfamilies showed different expression profiles in the antennae sensilla: SNMP1 proteins are detected in pheromone-sensitive olfactory receptor neurons (ORNs) [77]–[79]; however, the SNMP2 proteins are expressed in the supporting cells [78], [79]. In the present study, we have identified two SNMP genes, SNMP1 and SNMP2, in the A. ipsilon antennal transcriptomes (Table 5). Both have intact ORFs with lengths of 1569 bp and 1563 bp for SNMP1 and SNMP2, respectively, in agreement with our previous analyses [79]. The RT-PCR and RT-qPCR results revealed that both SNMP1 and SNMP2 were primarily expressed in the antennae of both sexes (Figure 7 and Figure 10). Furthermore, one gustatory receptor (AipsGR63) was identified in the A. ipsilon antennal transcriptomes (Table 5); AipsGR63 showed 46% amino acid identity with the B. mori gustatory receptor 63. The RT-PCR and RT-qPCR analyses showed that AipsGR63 was expressed in both the antennae and body part (Figure 7 and Figure 10).
Conclusions
Olfaction is an important sensory modality in insect. In present study we have successfully identified and annotated several groups of olfactory genes in the antennae of the noctuid moth A. ipsilon. The expression profile analysis revealed that 22 OBPs, 3 CSPs, 35 ORs, 14 IRs and the 2 SNMPs are uniquely or primarily expressed in the male and female antennae. These antennae-enriched OBPs, CSPs, ORs, IRs and SNMPs may play important physiological function in the pheromone and general odorant detection; thus, these genes could be meaningful targets for the study their biological functions, both in vivo and in vitro. An important direction of our future research will be the functional study of these olfactory genes.
Materials and Methods
Ethics statement
The black cutworm moth Agrotis ipsilon is common agricultural insect pests and are not included in the “List of Endangered and Protected Animals in China”. All operations were performed according to ethical guidelines in order to minimize pain and discomfort to the insects.
Insect rearing and tissue collection
The A. ipsilon colony was established in our laboratory in 2006. The larvae were reared with an artificial diet that was composed of wheat germ, casein and sucrose as the main components [63], [64], [79]. The laboratory colony was kept at 24°C with 75% relative humidity and a 16∶8 light:dark cycle. Pupae were sexed and maintained separately in hyaline plastic cups before emergence. Adult moths were given a 20% honey solution after emergence. Antennae were excised from 3-day-old male and female moths and immediately frozen and stored in liquid nitrogen until use.
RNA extraction and cDNA library construction
400 antennae from each sex were polled for total RNA extraction using TRIzol reagent using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) following the manufacturer’s instructions. The quantity of RNA samples was determined using a NanoDrop spectrophotometer (Thermo Scientific, Wilmington, DE, USA) and 1.1% agarose electrophoresis. Approximately 500 ng messenger RNA was further purified from 50 µg total RNA using a PolyATtract mRNA Isolation System III (Promega, Madison, WI, USA). The mRNAs were then sheared into approximately 800 nucleotides via RNA Fragmentation Solution (Autolab, Beijing, China) at 70°C for 30 sec, then cleaned and condensed using an RNeasy MinElute Cleanup Kit (Qiagen, Valencia, CA, USA). The first-strand cDNA was synthesized using N6 random primers and MMLV reverse transcriptase (TaKaRa, Dalian, China). Then, the second strand cDNAs were synthesized using secondary strand cDNA synthesis enzyme mixtures (Autolab, Beijing, China). The cDNAs with the desired length were purified using a QIAquick PCR Purification Kit (Qiagen, Valencia, CA, USA) and eluted with 10 µl elution buffer. After blunted and appended with a poly-A tail at the 3′ end according to Roche’s Rapid Library Preparing protocols (Roche, USA), the purified cDNAs were linked to GS-FLX Sequencing Adaptors (Roche, USA). Finally, the cDNAs that were shorter than 500 bp were removed using AMPure Beads according to the manufacture’s instructions (Beckman, USA) before the preparation of the cDNA library for next generation sequencing.
454 sequencing
Pyrosequencing of the cDNA library was performed by the Beijing Autolab Biotechnology Company using a 454 GS-FLX sequencer (Roche, IN, USA) according to the manufacturer’s instructions. All sequencing reads were deposited into the Short Read Archive (SRA) of the National Center for Biotechnology Information (NCBI), and can be accessed under the accession numbers SRR838973 and SRR838974 for the male and female antennal transcriptomes, respectively.
Sequence analysis and assembly
Base calling of the raw 454 reads in SFF files were performed using the python script sff_extract.py that was developed by COMAV (http://bioinf.comav.upv.es). All the raw reads were then processed to remove low quality and adaptor sequences using the programs TagDust [80], LUCY [81] and SeqClean [82] with default parameters. The resulting sequences were then screened against the NCBI UniVec database (http://www.ncbi.nlm.nih.gov/VecScreen/UniVec.html) to remove possible vector sequence contamination. The cleaned reads that were shorter than 60 bases were discarded based on the assumption that these reads might represent sequencing artifacts [83].
Two steps were taken to assemble the clean reads. First, the sequence assembler MIRA3 [84] was used with the assembly settings of a minimum sequence overlap of 30 bp and a minimum percentage overlap identity of 80%. Then, CAP3 was used with the assembly parameters of an overlap length cutoff >30 and an overlap percent identity cutoff >90% [85]. The resulting contigs and singletons that were more than 100 bases were retained as unigenes and annotated as described below.
Homology searches and functional classification
Following the assembly, homology searches of all unigenes were performed using the BLASTx and BLASTn programs against the GenBank non-redundant protein (nr) and nucleotide sequence (nt) databases at NCBI [86]. Matches with an E-value that was less than 1.0E-5 were considered significant [54]. Gene names were assigned to each unigene based on the best BLASTx hit with the highest score value.
Gene Ontology terms were assigned by the tool Blast2GO [87] through the BLASTx program with an E-value less than 1.0E-5. Then, the WEGO [88] software was used for the assignment of each GO ID to the related ontology entries. The longest open reading frame (ORF) of each unigene was determined by an ORF finder tool (http://www.ncbi.nlm.nih.gov/gorf/gorf.html).
Identification of A. ipsilon chemosensory genes
The tBLASTn program was performed, with available sequences of OBP, CSP, OR, GR, IR and SNMP proteins from Lepidoptera species as “query” to identify candidate unigenes encoding putative OBPs, CSPs, ORs, GRs, IRs and SNMPs in the A. ipsilon. All candidate OBPs, CSPs, ORs, GRs, IRs and SNMPs were manually checked by the BLASTx program at the National Center for Biotechnology Information (NCBI). The nucleotide sequences of all chemosensory genes that were identified from the A. ipsilon antennal transcriptomes are listed in Table S2.
Comparative analysis of chemosensory genes in the A. ipsilon male and female antennae
To compare the differential expression of chemosensory genes in the A. ipsilon male and female antennal transcriptomes, the read number for each chemosensory gene between male and female antennae was converted to RPKM (Reads Per Kilobase per Million mapped reads) [89], using the formula: RPKM (A) = (1,000,000×C×1,000)/(N×L), where RPKM (A) is the expression of chemosensory gene A, C is the number of reads that are uniquely aligned to chemosensory gene A, N is the total number of reads that are uniquely aligned to all unigenes, and L is the number of bases in chemosensory gene A. The FDR (false discovery rate) was used to determine the threshold of the P-value for multiple testing. FDR <0.001 and absolute values of the log2ratio >1 were used as the threshold to determine significant differences in gene expression. The RPKM method eliminates the influence of gene length and sequencing depth on the calculation of gene expression. Thus, the calculated gene expression can be directly used to compare gene expression between samples.
Sequence and phylogenetic analysis
The putative N-terminal signal peptides and the most likely cleavage site were predicted using the SignalP V3.0 program [90] (http://www.cbs.dtu.dk/services/SignalP/). Sequence alignments were performed using the program ClustalX 2.1 [91] with default gap penalty parameters of gap opening 10 and extension 0.2, and were edited using the GeneDoc 2.7.0 software. A neighbor-joining tree [92] was constructed using the program MEGA 5.0 [93] with a p-distance model and a pairwise deletion of gaps. The bootstrap support of tree branches was assessed by re-sampling amino acid positions 1000 times.
RT-PCR and RT-qPCR analysis
Two biological samples each with 150 male antennae, 150 female antennae and two moth body part (mixture of heads, thoraxes, abdomens, legs, wings) were used for RNA extraction using TRIzol reagent. Before transcription, total RNA was treated with RQ1 RNase-Free DNase (Promega, Madison, USA) to remove residual genomic DNA. cDNAs from male antennae, female antennae and the body part were synthesized using a GoScript Reverse Transcription System (Promega, Madison, USA). An equal amount of cDNA (200 ng) was used as RT-PCR and RT-qPCR templates. Specific primer pairs that were used for RT-PCR were designed with the program Primer 3 (http://frodo.wi.mit.edu/) (see Table S3). The β-actin (GenBank Acc. JQ822245) of A. ipsilon was used as the control gene to test the integrity of the cDNAs. The PCR was performed under following conditions: 95°C for 2 min, followed by 25–35 cycles (depending on the expression level of each gene) of 95°C for 30 sec, 56°C for 30 sec, 72°C for 1 min, and a final extension for 10 min at 72°C. PCR products were analyzed on 1.2% agarose gel and visualized after staining with ethidium bromide. To reach reproducibility, each sample was performed at least six times with two biological samples.
RT-qPCR analysis was conducted using an ABI 7500 Real-Time PCR System (Applied Biosystems, Carlsbad, CA). The primers that were used for RT-qPCR were designed using the program Beacon Designer 7.90 (PREMIER Biosoft International) (see Table S4). Two reference genes, β-actin (GenBank Acc. JQ822245) and ribosomal protein S3 (GenBank Acc. JQ822246) were used for normalizing the target gene expression and for correcting for sample-to-sample variation. Each RT-qPCR reaction was conducted in a 25 µl reaction mixture containing 12.5 µl of SuperReal PreMix Plus (TianGen, Beijing, China), 0.75 µl of each primer (10 µM), 0.5 µl of Rox Reference Dye, 1 µl of sample cDNA, and 9.5 µl of sterilized H2O. The RT-qPCR cycling parameters were as follows: 95°C for 15 min, followed by 40 cycles of 95°C for 10 sec and 60°C for 32 sec. Then, the PCR products were heated to 95°C for 15 sec, cooled to 60°C for 1 min, heated to 95°C for 30 sec and cooled to 60°C for 15 sec to measure the dissociation curves. Negative controls without either template or transcriptase were included in each experiment. To check reproducibility, each RT-qPCR reaction for each sample was performed in three technical replicates and two biological replicates. The comparative 2−ΔΔCT method [94] was used to calculate the relative quantification between tissues. The comparative analyses of each target gene among various tissues were determined using a one-way nested analysis of variance (ANOVA), followed by Tukey’s honestly significance difference (HSD) test using the software SPSS Statistics 18.0 (SPSS Inc., Chicago, IL, USA). When applicable, values were presented as the mean±SE.
Supporting Information
Table S1.
The 500 most highly abundant unigenes in the A. ipsilon antennal transcriptome.
https://doi.org/10.1371/journal.pone.0103420.s001
(XLSX)
Table S2.
The nucleotide sequences of 33 OBPs, 12 CSPs, 42 ORs, 24 IRs, 2 SNMPs and 1 GRs identified in present study.
https://doi.org/10.1371/journal.pone.0103420.s002
(DOCX)
Table S3.
Primers used for RT-PCR analysis of olfactory genes of the A. ipsilon moth.
https://doi.org/10.1371/journal.pone.0103420.s003
(DOCX)
Table S4.
Primers used for RT-qPCR analysis of olfactory genes of the A. ipsilon moth.
https://doi.org/10.1371/journal.pone.0103420.s004
(DOCX)
Table S5.
Protein names and sequences of ORs used in Figure 9.
https://doi.org/10.1371/journal.pone.0103420.s005
(TXT)
Table S6.
Protein names and sequences of IRs used in Figure 11.
https://doi.org/10.1371/journal.pone.0103420.s006
(TXT)
Acknowledgments
The authors thank Ms. Xiaoyang Qi and Wei Wang for their contributions in the insect rearing. The authors gratefully acknowledge the language help from Elsevier Language Editing Services.
Author Contributions
Conceived and designed the experiments: SHG YJZ JJZ. Analyzed the data: SHG KMW YYG XCL. Contributed reagents/materials/analysis tools: SHG. Contributed to the writing of the manuscript: SHG YJZ JJZ. Performed the sample collection, library construction, data processing, bioinformatics analysis and qRT-PCR: SHG. Performed the RT-PCR: LS RNY.
References
- 1. Kaissling KE (1986) Chemo-electrical transduction in insect olfaction receptors. Annu Rev Neurosci 9: 121–145.
- 2. Steinbrecht RA (1997) Pore structures in insect olfactory sensilla: a review of data and concepts. Int J Insect Morphol Embryol 26: 229–245.
- 3. Leal WS (2013) Odorant reception in insects: roles of receptors, binding proteins, and degrading enzymes. Annu Rev Entomol 58: 373–391.
- 4. Vogt RG, Riddiford LM (1981) Pheromone binding and inactivation by moth antennae. Nature 293: 161–163.
- 5. Klein U (1987) Sensillum-lymph proteins from antennal olfactory hairs of the moth Antheraea polyphemus (Saturniidae). Insect Biochem 17: 1193–1204.
- 6. Steinbrecht RA, Ozaki M, Ziegelberger G (1992) Immunocytochemical localization of pheromone-binding protein in moth antennae. Cell Tissue Res 270: 287–302.
- 7. Pelosi P (1996) Perireceptor events in olfaction. J Neurobiol 30: 3–19.
- 8. Xu P, Atkinson R, Jones DNM, Smith DP (2005) Drosophila OBP LUSH is required for activity of pheromone-sensitive neurons. Neuron 45: 193–200.
- 9. Laughlin JD, Ha TS, Jones DNM, Smith DP (2008) Activation of pheromone-sensitive neurons is mediated by conformational activation of pheromone-binding protein. Cell 133: 1255–1265.
- 10. Krieger MJ, Ross KG (2002) Identification of a major gene regulating complex social behavior. Science 295: 328–332.
- 11. McKenna MP, Hekmat-Scafe DS, Gaines P, Carlson JR (1994) Putative Drosophila pheromone-binding proteins expressed in a subregion of the olfactory system. J Biol Chem 269: 16340–16347.
- 12. Robertson HM, Martos R, Sears CR, Todres EZ, Walden KKO, et al. (1999) Diversity of odourant binding proteins revealed by an expressed sequence tag project on male Manduca sexta moth antennae. Insect Mol Biol 8: 501–518.
- 13. Angeli S, Ceron F, Scaloni A, Monti M, Monteforti G, et al. (1999) Purification, structural characterization, cloning and immunocytochemical localization of chemoreception proteins from Schistocerca gregaria. Eur J Biochem 262: 745–754.
- 14. Liu R, He X, Lehane S, Lehane M, Hertz-Fowler C, et al. (2012) Expression of chemosensory proteins in the tsetse fly Glossina morsitans morsitans is related to female host-seeking behavior. Insect Mol Biol 21: 41–48.
- 15. Jacquin-Joly E, Vogt RG, François MC, Nagnan-Le Meillour P (2001) Functional and expression pattern analysis of chemosensory proteins expressed in antennae and pheromonal gland of Mamestra brassicae. Chem Senses 26: 833–844.
- 16. González D, Zhao Q, McMahan C, Velasquez D, Haskins WE, et al. (2009) The major antennal chemosensory protein of red imported fire ant workers. Insect Mol Biol 18: 395–404.
- 17. Gu SH, Wang SY, Zhang XY, Ji P, Liu JT, et al. (2012) Functional characterizations of chemosensory proteins of the alfalfa plant bug Adelphocoris lineolatus indicate their involvement in host recognition. PLoS ONE 7(8): e42871.
- 18. Maleszka R, Stange G (1997) Molecular cloning, by a novel approach, of a cDNA encoding a putative olfactory protein in the labial palps of the moth Cactoblastis cactorum. Gene 202: 39–43.
- 19. Jin X, Brandazza A, Navarrini A, Ban L, Zhang S, et al. (2005) Expression and immunolocalisation of odorant-binding and chemosensory proteins in locusts. Cell Mol Life Sci 62: 1156–1166.
- 20. Nagnan-Le Meillour P, Cain AH, Jacquin-Joly E, François MC, Ramachandran S, et al. (2000) Chemosensory proteins from the proboscis of Mamestra brassicae. Chem Senses 25: 541–553.
- 21. Nomura A, Kawasaki K, Kubo T, Natori S (1992) Purification and localization of p10, a novel protein that increases in nymphal regenerating legs of Periplaneta americana (American cockroach). Int J Dev Biol 36: 391–398.
- 22. Kitabayashi AN, Arai T, Kubo T, Natori S (1998) Molecular cloning of cDNA for p10, a novel protein that increases in the regenerating legs of Periplaneta americana (American cockroach). Insect Biochem Mol Biol 28: 785–790.
- 23. Ban L, Scaloni A, Brandazza A, Angeli S, Zhang L, et al. (2003) Chemosensory proteins of Locusta migratoria. Insect Mol Biol 12: 125–134.
- 24. Zhou SH, Zhang J, Zhang SG, Zhang L (2008) Expression of chemosensory proteins in hairs on wings of Locusta migratoria (Orthoptera: Acrididae). J Appl Entomol 132: 439–450.
- 25. Guo W, Wang X, Ma Z, Xue L, Han J, et al. (2011) CSP and takeout genes modulate the switch between attraction and repulsion during behavioral phase change in the migratory locust. PLoS Genet 7(2): e1001291.
- 26. Sato K, Pellegrino M, Nakagawa T, Nakagawa T, Vosshall LB, et al. (2008) Insect olfactory receptors are heteromeric ligandgated ion channels. Nature 452: 1002–1006.
- 27. Wicher D, Schafer R, Bauernfeind R, Stensmyr MC, Heller R, et al. (2008) Drosophila odorant receptors are both ligand-gated and cyclic-nucleotide-activated cation channels. Nature 452: 1007–1011.
- 28. Nakagawa T, Sakurai T, Nishioka T, Touhara K (2005) Insect sex-pheromone signals mediated by specific combinations of olfactory receptors. Science 307: 1638–1642.
- 29. van der Goes van Naters W, Carlson JR (2007) Receptors and neurons for fly odors in Drosophila. Curr Biol 17: 606–612.
- 30. Clyne PJ, Warr CG, Freeman MR, Lessing D, Kim J, et al. (1999) A novel family of divergent seven transmembrane proteins: Candidate odorant receptors in Drosophila. Neuron 22: 327–338.
- 31. Gao Q, Chess A (1999) Identification of candidate Drosophila olfactory receptors from genomic DNA sequence. Genomics 60: 31–39.
- 32. Vosshall LB, Amrein H, Morozov PS, Rzhetsky A, Axel R (1999) A spatial map of olfactory receptor expression in the Drosophila antenna. Cell 96: 725–736.
- 33. Hill CA, Fox AN, Pitts RJ, Kent LB, Tan PL, et al. (2002) G protein-coupled receptors in Anopheles gambiae. Science 298: 176–178.
- 34. Bohbot J, Pitts RJ, Kwon HW, Rützler M, Robertson, et al (2007) Molecular characterization of the Aedes aegypti odorant receptor gene family. Insect Mol Biol 16: 525–537.
- 35. Robertson HM, Wanner KW (2006) The chemoreceptor superfamily in the honey bee, Apis mellifera: expansion of the odorant, but not gustatory, receptor family. Genome Res 16: 1395–1403.
- 36. Robertson HM, Gadau J, Wanner KW (2010) The insect chemoreceptor superfamily of the parasitoid jewel wasp Nasonia vitripennis. Insect Mol Biol 19 (Suppl. 1)121–36.
- 37. Wanner KW, Anderson AR, Trowell SC, Theilmann DA, Robertson HM, et al. (2007) Female-biased expression of odourant receptor genes in the adult antennae of the silkworm, Bombyx mori. Insect Mol Biol 16: 107–119.
- 38. Engsontia P, Sanderson AP, Cobb M, Walden KKO, Robertson HM, et al. (2008) The red flour beetle’s large nose: an expanded odorant receptor gene family in Tribolium castaneum. Insect Biochem Mol Biol 38: 38–397.
- 39. Smadja C, Shi P, Butlin RK, Robertson HM (2009) Large gene family expansions and adaptive evolution for odorant and gustatory receptors in the pea aphid, Acyrthosiphon pisum. Mol Biol Evol 26: 2073–2086.
- 40. Benton R, Vannice KS, Gomez-Diaz C, Vosshall LB (2009) Variant ionotropic glutamate receptors as chemosensory receptors in Drosophila. Cell 136: 149–162.
- 41. Croset V, Rytz R, Cummins SF, Budd A, Brawand D, et al. (2010) Ancient protostome origin of chemosensory ionotropic glutamate receptors and the evolution of insect taste and olfaction. PLoS Genet 6(8): e1001064.
- 42. Olivier V, Monsempes C, François MC, Poivet E, Jacquin-Joly E (2011) Candidate chemosensory ionotropic receptors in a Lepidoptera. Insect Mol Biol 20: 189–199.
- 43. Pitts RJ, Rinker DC, Jones PL, Rokas A, Zwiebel LJ (2011) Transcriptome profiling of chemosensory appendages in the malaria vector Anopheles gambiae reveals tissue- and sex-specific signatures of odor coding. BMC Genomics 12: 271.
- 44. Grosse-Wilde E, Kuebler LS, Bucks S, Vogel H, Wicher D, et al. (2011) Antennal transcriptome of Manduca sexta. Proc Natl Acad Sci USA 108: 7449–7454.
- 45. Bengtsson JM, Trona F, Montagné N, Anfora G, Ignell R, et al. (2012) Putative chemosensory receptors of the codling moth, Cydia pomonella, identified by antennal transcriptome analysis. PLoS ONE 7(2): e31620.
- 46. Liu Y, Gu S, Zhang Y, Guo Y, Wang G (2012) Candidate olfaction genes identified within the Helicoverpa armigera antennal transcriptome. PLoS ONE 7(10): e48260.
- 47. Nishimura O, Brillada C, Yazawa S, Maffei ME, Arimura G (2012) Transcriptome pyrosequencing of the parasitoid wasp Cotesia vestalis: genes involved in the antennal odorant-sensory system. PLoS ONE 7(11): e50664.
- 48. Mamidala P, Wijeratne AJ, Wijeratne S, Poland T, Qazi SS, et al. (2013) Identification of odor-processing genes in the emerald ash borer, Agrilus planipennis. PLoS ONE 8(2): e56555.
- 49. Gu SH, Wu KM, Guo YY, Field LM, Pickett JA, et al. (2013) Identification and expression profiling of odorant binding proteins and chemosensory proteins between two wingless morphs and a winged morph of the cotton aphid Aphis gossypii Glover. PLoS ONE 8(9): e73524.
- 50. Poivet E, Gallot A, Montagné N, Glaser N, Legeai F, et al. (2013) A comparison of the olfactory gene repertoires of adults and larvae in the noctuid moth Spodoptera littoralis. PLoS ONE 8(4): e60263.
- 51. Andersson MN, Grosse-Wilde E, Keeling CI, Bengtsson JM, Yuen MM, et al. (2013) Antennal transcriptome analysis of the chemosensory gene families in the tree killing bark beetles, Ips typographus and Dendroctonus ponderosae (Coleoptera: Curculionidae: Scolytinae). BMC Genomics 14: 198.
- 52. Rings RW, Arnold FJ, Keaster AJ, Musick GJ (1974) A worldwide, annotated bibliography of the black cutworm, Agrotis ipsilon (Hufnagel). Ohio Agric Res Dev Cent Res Circ 198: 1–106.
- 53. Clement SL, Show ED, Way MO (1982) Black cutworm pheromone trapping in strawberries. Calif Agric 36: 20–21.
- 54. Anderson I, Brass A (1998) Searching DNA databases for similarities to DNA sequences: when is a match significant? Bioinformatics14: 349–356.
- 55. Jacquin-Joly E, Legeai F, Montagné N, Monsempes C, François MC, et al. (2012) Candidate chemosensory genes in female antennae of the noctuid moth Spodoptera littoralis. Int J Biol Sci 8: 1036–1050.
- 56. Ashburner M, Ball CA, Blake JA, Botstein D, Butler H, et al. (2000) Gene ontology: tool for the unification of biology. Nat Genet 25: 25–29.
- 57. Pelosi P, Zhou JJ, Ban LP, Calvello M (2006) Soluble proteins in insect chemical communication. Cell Mol Life Sci 63: 1658–1676.
- 58. Leal WS (2005) Pheromone reception. Top Curr Chem 240: 1–36.
- 59. Zhou JJ (2010) Odorant-binding proteins in insects. Vitam Horm 83: 241–272.
- 60. Gong DP, Zhang HJ, Zhao P, Xia QY, Xiang ZH (2009) The odorant binding protein gene family from the genome of silkworm, Bombyx mori. BMC Genomics 10: 332.
- 61. Xu PX, Zwiebel LJ, Smith DP (2003) Identification of a distinct family of genes encoding atypical odorant-binding proteins in the malaria vector mosquito, Anopheles gambiae. Insect Mol Biol 12: 549–560.
- 62. Hekmat-Scafe DS, Scafe CR, McKinney AJ, Tanouye MA (2002) Genome-wide analysis of the odorant-binding protein gene family in Drosophila melanogaster. Genome Res 12: 1357–1369.
- 63. Gu SH, Zhou JJ, Wang GR, Zhang YJ, Guo YY (2013) Sex pheromone recognition and immunolocalization of three pheromone binding proteins in the black cutworm moth Agrotis ipsilon. Insect Biochem Mol Biol 43: 237–251.
- 64. Gu SH, Wu KM, Guo YY, Pickett JA, Field LM, et al. (2013) Identification of genes expressed in the sex pheromone gland of the black cutworm Agrotis ipsilon with putative roles in sex pheromone biosynthesis and transport. BMC Genomics 14: 636.
- 65. Wanner KW, Willis LG, Theilmann DA, Isman MB, Feng QL, et al. (2004) Analysis of the insect os-d-like gene family. J Chem Ecol 30: 889–911.
- 66. Vogel H, Heidel AJ, Heckel DG, Groot AT (2010) Transcriptome analysis of the sex pheromone gland of the noctuid moth Heliothis virescens. BMC Genomics 11: 29.
- 67. Gong DP, Zhang HJ, Zhao P, Lin Y, Xia QY, et al. (2007) Identification and expression pattern of the chemosensory protein gene family in the silkworm, Bombyx mori. Insect Biochem Mol Biol 37: 266–277.
- 68. Dani FR, Michelucci E, Francese S, Mastrobuoni G, Cappellozza S, et al. (2011) Odorant-binding proteins and chemosensory proteins in pheromone detection and release in the silkmoth Bombyx mori. Chem Senses 36: 335–344.
- 69. Schneider D, Schulz S, Priesner E, Ziesmann J, Francke W (1998) Autodetection and chemistry of female and male pheromone in both sexes of the tiger moth Panaxia quadripunctaria. J Comp Physiol A 182: 153–161.
- 70. Krieger J, Grosse-Wilde E, Gohl T, Dewer YME, Raming K, et al. (2004) Genes encoding candidate pheromone receptors in a moth (Heliothis virescens). Proc Natl Acad Sci USA 101: 11845–11850.
- 71. Greiner B, Gadenne C, Anton S (2004) Three-dimensional antennal lobe atlas of the male moth, Agrotis ipsilon: A tool to study structure–function correlation. J Comp Neurol 475: 202–210.
- 72. Carlsson MA, Galizia CG, Hansson BS (2002) Spatial representation of odours in the antennal lobe of the moth Spodoptera littoralis (Lepidoptera: Noctuidae). Chem Senses 27: 231–244.
- 73. Vosshall LB (2001) The molecular logic of olfaction in Drosophila. Chem Senses 26: 207–213.
- 74. Benton R, Vannice KS, Vosshall LB (2007) An essential role for a CD36-related receptor in pheromone detection in Drosophila. Nature 450: 289–293.
- 75. Jin X, Ha TS, Smith DP (2008) SNMP is a signaling component required for pheromone sensitivity in Drosophila. Proc Natl Acad Sci USA 105: 10996–11001.
- 76. Vogt RG, Miller NE, Litvack R, Fandino RA, Sparks J, et al. (2009) The insect SNMP gene family. Insect Biochem Mol Biol 39: 448–456.
- 77. Rogers ME, Steinbrecht RA, Vogt RG (2001) Expression of SNMP-1 in olfactory neurons and sensilla of male and female antennae of the silkmoth Antheraea polyphemus. Cell Tissue Res 303: 433–446.
- 78. Forstner M, Gohl T, Gondesen I, Raming K, Breer H, et al. (2008) Differential expression of SNMP-1 and SNMP-2 proteins in pheromone-sensitive hairs of moths. Chem Senses 33: 291–299.
- 79. Gu SH, Yang RN, Guo MB, Wang GR, Wu KM, et al. (2013) Molecular identification and differential expression of sensory neuron membrane proteins in the antennae of the black cutworm moth Agrotis ipsilon. J Insect Physiol 59: 430–443.
- 80. Lassmann T, Hayashizaki Y, Daub CO (2009) TagDust–a program to eliminate artifacts from next generation sequencing data. Bioinformatics 25: 2839–2840.
- 81. Chou HH, Holmes MH (2001) DNA sequence quality trimming and vector removal. Bioinformatics 17: 1093–1104.
- 82. Chen YA, Lin CC, Wang CD, Wu HB, Hwang PI (2007) An optimized procedure greatly improves EST vector contamination removal. BMC Genomics 8: 416.
- 83. Meyer E, Aglyamova GV, Wang S, Buchanan-Carter J, Abrego D, et al. (2009) Sequencing and de novo analysis of a coral larval transcriptome using 454 GSFlx. BMC Genomics 10: 219.
- 84. Chevreux B, Pfisterer T, Drescher B, Driesel AJ, Müller WEG, et al. (2004) Using the miraEST assembler for reliable and automated mRNA transcript assembly and SNP detection in sequenced ESTs. Genome Res 14: 1147–1159.
- 85. Huang X, Madan A (1999) CAP3: A DNA sequence assembly program. Genome Res 9: 868–877.
- 86. Altschul SF, Madden TL, Schäffer AA, Zhang JH, Zhang Z, et al. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25: 3389–3402.
- 87. Conesa A, Götz S, García-Gómez JM, Terol J, Talón M, et al. (2005) Blast2GO: a universal tool for annotation, visualization and analysis in functional genomics research. Bioinformatics 21: 3674–3676.
- 88. Ye J, Fang L, Zheng HK, Zhang Y, Chen J, et al. (2006) WEGO: a web tool for plotting GO annotations. Nucleic Acids Res 34: W293–W297.
- 89. Mortazavi A, Williams BA, McCue K, Schaeffer L, Wold B (2008) Mapping and quantifying mammalian transcriptomes by RNA-Seq. Nat Methods 5: 621–628.
- 90. Bendtsen JD, Nielsen H, von Heijne G, Brunak S (2004) Improved prediction of signal peptides: SignalP 3.0. J Mol Biol 340: 783–795.
- 91. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25: 4876–4882.
- 92. Saitou N, Nei M (1987) The neighbor-joining method: A new method for reconstructing phylogenetic trees. Mol Biol Evol 4: 406–425.
- 93. Tamura K, Dudley J, Nei M, Kumar S (2007) MEGA4: Molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol Biol Evol 24: 1596–1599.
- 94. Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCt method. Methods 25: 402–408.